Tomato Genetic System
The genus Lycopersicon includes the cultivated tomato (L. esculentum Mill.) together with its wild relatives. The wild species bear a wealth of genetic variability. Less than 10% of the total genetic diversity in the Lycopersicon gene pool is found in L. esculentum (Miller and Tanksley, 1990). The center of diversity for tomato is located in western South America, and the cherry tomato L. esculentum var. cerasiforme is considered as the most likely ancestor of cultivated tomatoes. Karyotypes of the Lycopersicon species are very similar with little or no structural difference among species (Barton, 1950). As a crop plant, tomato is one of the best-characterized plant systems. It has a relatively small genome of 0.95 pg or 950 Mb per haploid nucleus, (Arumuganathan and Earle, 1991) and features such as diploidy, self pollination, and a relatively short generation time make it amenable to genetic analysis. Classical genetics has created one of the largest stocks of morphological mutations induced by radiation (X-rays, UV-light, neutrons) and chemical mutagenesis. A major contributor in the mutagenesis area was Hans Stubbe who developed over 300 L. esculentum mutants and 200 in L. pimpinellifolium. (for summary see Rick, 1975). A particularly interesting example of induced mutagenesis was the directed manipulation of fruit size of L. esculentum and L. pimpinellifolium.(Stubbe, 1971).
A considerable proportion of these mutations have been mapped onto the classical genetic map. By 1988, the classical linkage map of the tomato genome comprised of 233 morphological and isozyme loci. An additional 86 have been assigned to their respective chromosomes via two-point or trisomic tests. The number of mapped genes in the form of cDNAs has increased considerably with the introduction of RFLP markers. The current tomato RFLP map was constructed using an F2 population of the interspecific cross L. esculentum x L. pennellii and contains more than 1030 markers, which were distributed over 1276 cM. (Tanksley et al., 1992). A number of morphological and isozyme markers have also been mapped with respect to RFLP markers orienting the molecular linkage map with both the classical morphological and cytological maps of tomato. An integrated high-density RFLP-AFLP map of tomato based on two independent L. esculentum x L. pennellii F2 populations has been constructed (Haanstra et al., 1999). This map spanned 1482 cM and contained 67 RFLP and 1175 AFLP markers. Both RFLP and AFLP maps show clusters of markers associated with almost all centromeres and some telomeres indicating that recombination is suppressed in those regions.
The current tomato map is considered to be complete in that all molecular and classical markers could be mapped to one of the 12 linkage groups indicating that no loci failed to link up with the map. The average relationship between genetic and physical distance in tomato is about 750 kb per cM. The actual ratio of genetic and physical distance varies considerably depending on the chromosomal region. High-resolution genetic and physical mapping around the Tm-2a region, which is located close to the centromere of chromosome 9, indicates that one cM in this area corresponds to more than five million base pairs (Pillen et al., 1996), approximately a sevenfold suppression of recombination over the expected value based on the estimated physical size of the region. In contrast, map-based cloning of the chloronerva gene, which is involved in iron uptake and located in euchromatin of chromosome 1, demonstrated that the ratio of genetic to physical distance in the chloronerva region is 160 kb per 1 cM (Ling et al., 1999) suggesting much higher levels of recombination in this area of the genome. By determining frequency and distribution of recombination nodules on tomato synaptonemal complexes, Sherman and Stack (1995) observed a much lower frequency of recombination nodules in heterochromatic regions around the centromeres compared to euchromatin. Suppression of recombination near the centromeres and higher values of recombination in distal chromosomal regions were also observed in potato (Bonierbale, 1988; Tanksley et al., 1992) and many other plant and animal species. The tomato genome at the DNA level is comprised of approximately 78% single copy sequences, as evaluated under high stringency hybridization conditions (Zamir and Tanksley, 1988). In other plant species with large genome sizes, such as wheat or pea, the single copy fraction is less than 20%, and in barley and rye, it is less than 50%. The remaining part of the tomato sequences is repetitive DNA of which four major classes have been characterized. Ribosomal DNA represents the most abundant repetitive DNA family and comprises approximately 3% of the tomato genome. Both 5S and 45S rRNA genes are tandemly repeated with 1,000 and 2,300 copies and map to single loci on chromosome 1 and 2, (Vallejos et al., 1986; Lapitan et al., 1991) respectively. As confirmed by in situ hybridization, a 162 bp tandem repeat, TGRI, with 77,000 copies in the genome is localized within a few hundred kb of the terminal 7 bp telomeric repeat TT(T/A)AGGG at 20 of 24 chromosome ends; (Ganal et al., 1988) and, in addition, it is also found at a few centromeric and interstitial sites (Lapitan et al., 1989; Ganal et al., 1992). Two other tomato genomic repeats, TGRII and TGRIII, are less abundant with 4,200 and 2,100 copies, respectively. TGRII is apparently randomly distributed with an average spacing of 133 kb, and TGRIII is predominantly clustered in the centromeric regions of chromosomes. Except TGRIII, these repeats are only present in Lycopersicon species. (Ganal et al., 1988).
Zamir and Tanksley (1988) also reported a positive correlation between copy number and rate of divergence of repeats among DNA sequences from related solanaceous species. The more highly repeated sequences evolve more rapidly, whereas single copy coding regions are more conserved among different species. 43% of cloned low copy telomere-homologous sequences, which were mapped near the tomato centromeres, hybridized to DNA from L. esculentum but not to L. pennellii, whereas single copy probes hybridized to both L. esculentum and L. pennellii (Presting et al., 1996) indicating rapid evolution of centromere-proximal sequences. Cytologically, the centromere of higher eukaryotes is a constriction on condensed metaphase chromosomes surrounded by large blocks of pericentric heterochromatin. At the primary constriction, various proteins associate with the centromeric DNA and form the kinetochore, the attachment point for the spindle apparatus. Thus, centromeres composed of both DNA sequences and proteins organized in a structurally and functionally unique manner and are complex genetic loci. Kinetochore protein components appear to be more conserved than centromeric sequences. As a general rule, plant centromeric DNA is heterogenous, composed of megabases of satellite DNA with poor conservation of primary repeat sequence across distantly related plant species, and includes also low-copy sequences, transposable elements and telomere-similar repeats. Each tomato chromosome has heterochromatin concentrated around its centromere. Using Feulgen densitometry and SC karyotype data, it was determined that 77% of the DNA in tomato pachytene chromosomes is packaged in heterochromatin, which is similar to an earlier estimate (75.3%) in mitotic metaphase chromosomes (Peterson et al., 1996). In association with findings of Zamir and Tanksley (1988), these data suggest that a large fraction of tomato heterochromatic DNA is composed of single- and/or low-copy sequences and makes tomato heterochromatin unusual and probably genetically active.
approximate map position of the centromere is now known for each tomato
chromosome. For chromosomes 1 and 2, the centromere positions have been
identified by RFLP mapping and by in situ hybridization with 5S rDNA and
45S rDNA (Lapitan et al., 1991; Tanksley et al., 1988), respectively.
The centromeres of chromosomes 3 and 6 have been located on the integrated
molecular-classical map and by deletion mapping (Van-der Biezen et al.,
1994; Van Wordragen et al., 1994). Since there is evidence that the potato/tomato
inversions on chromosomes 5, 10, 11 and 12 involve entire chromosome arms,
the respective centromeres are most likely located at the inversion breakpoints
(Tanksley et al., 1992). Map positions of the centromeres of chromosomes
4 and 8 were predicted based on the relationship among the cytological,
genetic and molecular tomato maps. RFLP hybridization and dosage analysis
of telo-, secondary and tertiary trisomic stocks (Frary et al., 1996)
have achieved a more precise localization of the centromeres of chromosomes
7 and 9. Despite their functional importance, the molecular characteristics
of the centromeres of higher eukaryotes remain ill-defined. The most extensively
studied DNA sequence is the 171 bp alpha satellite sequence, which is
located exclusively at the primary constriction of human chromosomes and
thought to play a major structural and/or functional role at human centromeres
(Haaf et al., 1992; Harrington et al., 1997). So far, no plant DNA sequences
essential for centromere function have been identified. The only plant
sequence to which a centromere function could be attributed is a DNA repeat
from the centric region of the maize B chromosome. Sequence analysis revealed
that this repeat contains several motifs. One is a stretch of repeats,
which has high similarity to the telomeric repeat of plants. The other
has a significant homology to the 180 bp repeat that comprises the telomeric
heterochromatic knob. Under certain conditions such knobs can function
as spindle attachment sites and form neocentromeres (Alfenito and Bichler,
1993). Two repetitive sequences CentA and CentC were characterized in
the centromeric region of the maize chromosome 9 (Ananiev et al., 1998).
CentA has a structural similarity to retroelements and CentC is a tandem
repeat which forms clusters of different sizes at centromeric sites of
all maize chromosomes without obvious homology to the maize knob-associated
tandem repeat. Several types of DNA sequences located at pericentromeric
regions have been identified, but their role in centromere function remains
elusive. (Martinez-Zapater et al., 1986; Ganal et al., 1988; Maluszynska
and Heslop-Harrison, 1991; Richards et al., 1991; Aragon-Alcaide et al.,
1996; Jiang et al., 1996; Presting et al., 1996; Thompson et al., 1996).
Centromere research has a potentally important application in the production
of artificial chromosomes for use as plant cloning vectors. Studying of
plant centromeric DNA sequences provides an opportunity to look for conserved
structural patterns or primary nucleotide sequence motifs that may contribute
to centromere function. Recently, genome mapping interest has been directed
to the location of microsatellite sequences on tomato chromosomes. Microsatellite
polymorphism and genomic distribution were studied by fingerprinting of
the tomato genome using labeled oligonucleotide probes complementary to
GATA or GACA microsatellites. The copy number and the size of microsatellite
containing restriction fragments were highly variable between tomato cultivars
(Vosman et al., 1992). The mapping of individual fingerprint bands containing
GATA or GACA microsatellites showed predominant association of these repeats
with tomato centromeres. Structure, abundance, variability and location
were evaluated for a number of different simple sequence repeats isolated
from genomic libraries (Broun and Tanksley, 1996). Ten generated microsatellite
markers (6(GT)n, 3(GA)n and 1(ATT)n) were tested for polymorphism in a
set of ten tomato cultivars. Only two microsatellite loci ((GA)16 and
an imperfect ATT repeat) did reveal significant polymorphism in the tested
cultivars and were mapped on the high resolution molecular map near the
putative centromeres of chromosomes 3 and 12, respectively. In addition,
nine polymorphic GATA-containing RFLP fragments (9-15 kb) were scored
as dominant markers and mapped within clusters of markers adjacent to
Miller, J.C., and S.D. Tanksley.1990. RFLP analysis of phylogenetic relationships
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Prepared by M. Abhary (firstname.lastname@example.org)
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